ORIGINAL PAPER

The copper centers of tyramine b-monooxygenaseand its catalytic-site methionine variants:an X-ray absorption study

Corinna R. Hess • Judith P. Klinman •

Ninian J. Blackburn

Received: 1 April 2010 / Accepted: 12 May 2010 / Published online: 11 June 2010

� The Author(s) 2010. This article is published with open access at Springerlink.com

Abstract Tyramine b-monooxygenase (TBM) is a

member of a family of copper monooxygenases containing

two noncoupled copper centers, and includes peptidylgly-

cine monooxygenase and dopamine b-monooxygenase. In

its Cu(II) form, TBM is coordinated by two to three His

residues and one to two non-His O/N ligands consistent

with a [CuM(His)2(OH2)2–CuH(His)3(OH2)] formulation.

Reduction to the Cu(I) state causes a change in the X-ray

absorption spectroscopy (XAS) spectrum, consistent with a

change to a [CuM(His)2S(Met)–CuH(His)3] environment.

Lowering the pH to 4.0 results in a large increase in the

intensity of the Cu(I)–S extended X-ray absorption fine

structure (EXAFS) component, suggesting a tighter Cu–S

bond or the coordination of an additional sulfur donor. The

XAS spectra of three variants, where the CuM Met471

residue had been mutated to His, Cys, and Asp, were

examined. Significant differences from the wild-type

enzyme are evident in the spectra of the reduced mutants.

Although the side chains of His, Cys, and Asp are expected

to substitute for Met at the CuM site, the data showed

identical spectra for all three reduced variants, with no

evidence for coordination of residue 471. Rather, the K-

edge data suggested a modest decrease in coordination

number, whereas the EXAFS indicated an average of two

His residues at each Cu(I) center. These data highlight the

unique role of the Met residue at the CuM center, and pose

interesting questions as to why replacement by the cupro-

philic thiolate ligand leads to detectable activity whereas

replacement by imidazole generates inactive TBM.

Keywords Copper � Tyramine b-monooxygenase �X-ray absorption � Extended X-ray absorption fine structure

Abbreviations

DBM Dopamine b-monooxygenase

EXAFS Extended X-ray absorption fine structure

FT Fourier transform

PHM Peptidylglycine monooxygenase

S2 Schneider 2

TBM Tyramine b-monooxygenase

Tris Tris(hydroxymethyl)aminomethane

XANES X-ray absorption near edge structure

XAS X-ray absorption spectroscopy

Introduction

Tyramine b-monooxygenase (TBM) is a member of a

small class of copper-containing monooxygenases which

Electronic supplementary material The online version of thisarticle (doi:10.1007/s00775-010-0677-3) contains supplementarymaterial, which is available to authorized users.

C. R. Hess � J. P. Klinman

Department of Chemistry and of Molecular and Cell Biology,

University of California,

Berkeley, CA 94720, USA

N. J. Blackburn (&)

Department of Science and Engineering,

School of Medicine,

Oregon Health and Sciences University,

Beaverton, OR 97006, USA

e-mail: ninian@comcast.net

C. R. Hess

Chemistry Department,

Durham University,

Durham DH1 3LE, UK

123

J Biol Inorg Chem (2010) 15:1195–1207

DOI 10.1007/s00775-010-0677-3

utilize a pair of mononuclear copper centers at their active

sites [1–4] to catalyze the hydroxylation of benzylic or

peptidylglycyl Ca carbon atoms. Other members of the

family include dopamine b-monooxygenase (DBM; cate-

cholamine biosynthesis), peptidylglycine monooxygenase

(PHM; C-terminal peptide amidation), and monooxygenase

X (unknown function) [5]. Akin to the mammalian

enzymes DBM and PHM, TBM is associated with neuro-

transmission in invertebrates, and catalyzes the hydroxyl-

ation of tyramine to octopamine (Fig. 1b); both molecules

are critical to physiological functions, such as neuromus-

cular transmission and behavioral development, in insects

[6–8]. Whereas a significant database of spectroscopic

information has been accumulated for these enzymes

[9–13], crystallographic characterization has only been

achieved for PHM [14, 15]. The two copper centers of the

monooxygenases (denoted CuM and CuH) are bound in a

solvent-filled cleft approximately 11 A apart, and are

structurally and spectroscopically distinct. CuM is coordi-

nated by two His residues and a Met residue, whereas CuH

is coordinated to three His residues. A structure of the

reduced enzyme co-crystallized with a slow substrate has

revealed the presence of a dioxygen molecule bound at the

CuM center, where the O–O bond length is suggestive of a

Cu(II)–superoxo complex [16]. This observation, together

with further kinetic [17, 18], biochemical [19], and theo-

retical [20] evidence, has led to the proposal that the active

oxygen species is best formulated as a Cu(II)–superoxo

species.

The Met residue at the CuM center appears to play a

critical but as yet elusive role in catalysis. Early studies on

PHM M314X substitutions reported undetectable catalytic

activity in spent medium from PHM cell lines expressing

the Ile, His, Asp, and Cys variants [21, 22]. More recent

studies using purified variants of TBM in which the

homologous Met ligand (M471) was mutated to His, Asp,

or Cys have established a dramatic decrease in activity,

with only the M471C variant displaying measurable

activity [9]. Extended X-ray absorption fine structure

(EXAFS) studies on both DBM [10, 11] and PHM [12, 23,

24] have determined that the sulfur from the Met residue is

a ligand to CuM in the reduced but not the oxidized

enzyme. However, a crystal structure of the M314I mutant

of PHM revealed changes at both CuM and CuH suggestive

of a more complex role than simple copper coordination

[25]. Model studies on mononuclear Cu(II)–superoxo

complexes have in general provided limited insight into the

contribution of a coordinated thioether to oxygen activation

[26]. The thioether moiety has little effect on the reactivity

of the synthetic copper complexes. However, incorporation

of thioether into a mononuclear b-diketonate Cu(II)

superoxo species suggested complex chemistry involving

fluxional Met coordination at low temperature [27, 28],

which complemented observations from EXAFS that the

PHM Cu–S(Met) bond is unusually weak, and also possibly

fluxional. This led to the suggestion that the dynamics of

the Cu–S(Met) interaction in PHM may play a role in

providing a vibrational coordinate for hydrogen-atom tun-

neling during hydrogen-atom abstraction from the peptide

substrate [24].

Other explanations for the important role of the M-site

Met residue in catalysis have been advanced. It has been

suggested that the weak donor power of the Met residue

[17] prevents the copper–dioxygen complex from under-

going significant O2 reduction prior to substrate activation

by hydrogen-atom abstraction. In essence, this effect

would shift the equilibrium for Cu(I)–dioxygen binding to

the left, by S(Met) stabilization of the Cu(I) form, thereby

inhibiting uncoupling reactions induced by superoxide or

peroxide ‘‘leakage’’ from the active site [17, 20]. A Cu(I)

stabilizing role of Met has been observed experimentally

in studies of the peroxide reactivity of PHM where

analysis of product isotopomers clearly showed an equi-

librium between Cu(II)–superoxo and Cu(I)–dioxygen

species [19]. However, despite much work, the role of the

thioether ligand in catalysis remains a major unanswered

question.

The expression of TBM in Drosophila Schneider 2 (S2)

cells [4, 29] provides a useful platform for mutagenesis

studies to further probe the role of the Met residue at CuM

in catalysis. Recently copper binding and enzymatic

activity data were reported for a series of mutants at the

M471 residue in TBM, including the Met to Cys, His, and

Asp variants [9]. As stated previously, only the Cys variant

retained measurable catalytic activity. The oxidized Cu(II)

forms of all three variants bound an equivalent amount of

copper, and showed no perturbation of their EPR spectral

properties relative to the wild-type protein. In the present

study, EXAFS measurements on these isolated TBM

mutants have allowed us to examine, for the first time,

potential structural correlations to the altered reactivities.

Since the Met residue forms only a weak axial interaction

in the oxidized form of PHM [13, 23] and DBM [10],

major differences in copper coordination are only antici-

pated in the reduced forms. Here, we used X-ray absorption

spectroscopy (XAS) to probe the structure of the copper

centers in the wild type and each of the M471 variants, and

in particular to assess whether residue 471 is able to bind to

the reduced copper center. Changes in the Cu(I) coordi-

nation sphere of the mutant enzyme forms would have

significant implications for the reaction of the enzyme with

dioxygen, and for the subsequent formation of relevant

intermediates. Although wild-type TBM appears to bind

M471 in a fashion similar to PHM and DBM, we find no

spectroscopic evidence for strong interactions of His, Asp,

or Cys with the CuM center.

1196 J Biol Inorg Chem (2010) 15:1195–1207

123

Materials and methods

Materials

Drosophila S2 cells, insect cell growth media, and Dro-

sophila Expression System were purchased from Invitro-

gen. Blasticidin was purchased from Sigma. Primers were

custom-ordered, high-performance-liquid-chromatography-

purified, from Operon. Chromatography media was

purchased from GE Healthcare. All other materials were

obtained from Sigma.

Protein expression

For the purposes of the XAS studies, wild-type and mutant

TBM lacking the His-tag were used. A stop codon was

introduced into the pBipTBM plasmid DNA for wild-type

TBM and TBM mutants [9], preceding the sequence of bases

encoding for the rTEV recognition site and His-tag

contained in the original construct [4]. The altered

pBipTBM plasmid was transformed into Escherichia coli

strain XL1 Blue (Stratagene) cells and purified using a

QIAGEN HiSpeed plasmid midi kit. The composition of the

purified plasmid was confirmed by DNA sequencing (Uni-

versity of California, Berkeley, Sequencing Facility), prior

to transfection into the S2 cells. The expression of wild-type

and mutant TBM in Drosophila S2 cells was performed

according to previously described procedures [4].

Protein purification

All purification steps were carried out at 277 K. The

recombinant enzyme was purified from the culture medium

as follows. The cell culture (1.5 L) was centrifuged

(3,000 rpm, 10 min) and the supernatant batch-bound to

200 mL (diethylamino)ethyl–Sepharose Fast Flow resin

(10 mM potassium phosphate, pH 8), as previously

described [4]. The column-bound protein was washed with

OHHH

NH2

OHOHH

NH2

TBM

His Asp Cys

(a)

(b)

(c)

Fig. 1 a The H and M centers

of the mononuclear

monooxygenases modeled on

the crystal structure (Protein

Data Bank file 3PHM) of the

catalytic core of peptidylglycine

monooxygenase. b Reaction

catalyzed by tyramine b-

monooxygenase. c Modeling of

His, Asp, and Cys at residue 471

into the catalytic M-site

showing the potential for

coordination of each ligand at

the CuM center

J Biol Inorg Chem (2010) 15:1195–1207 1197

123

2 L of equilibration buffer, and the protein was subse-

quently eluted with a buffer gradient (1 L, 10 mM to

0.25 M potassium phosphate, pH 8). Protein-containing

fractions (as determined by sodium dodecyl sulfate poly-

acrylamide gel electrophoresis) were combined, dialyzed

overnight (10 kDa cutoff membrane) against 4 L of

tris(hydroxymethyl)aminomethane (Tris) buffer (20 mM

Tris, 50 mM NaCl, pH 7.5), concentrated to approximately

20 mL (Millipore Ultrafree centrifugal concentrators,

50 kDa cutoff membrane), and loaded onto a Q-Sepharose

column (15 mL) equilibrated with the same low-salt buffer.

The column was washed with 100 mL of the equilibration

buffer, and the protein was eluted using a NaCl gradient

(300 mL, 20 mM Tris, 0.05 M to 0.2 M NaCl; followed by

100 mL, 0.2 M to 0.5 M NaCl). The protein content of the

fractions was again determined by sodium dodecyl sulfate

polyacrylamide gel electrophoresis. The purest fractions

were combined, concentrated to less than 1 mL, and puri-

fied using size-exclusion chromatography, as described

previously [4]. Single-banded fractions were combined and

used for the preparation of XAS samples.

Preparation of samples for XAS

The purified wild-type and mutant TBM samples (3–6 mg,

50 mM Tris, 0.1 M NaCl, pH 7.5) were copper-reconsti-

tuted by dialysis against a tenfold volume of the same Tris

buffer containing 40 lM CuSO4, for 6 h (10 kDa cutoff

membrane). The samples were subsequently concentrated

to 1 mL (10 kDa cutoff membrane), diluted to 5 mL with

low-copper-containing buffer (50 mM Tris, 0.1 M NaCl,

5 lM CuSO4, 24% ethylene glycol, pH 7.5), and finally

reconcentrated to a volume of less than 1 mL.

The low-pH wild-type TBM sample was prepared by

12-fold dilution of a concentrated sample at pH 7.5 with

low-pH Tris buffer (50 mM Tris, 0.1 M NaCl, 21% eth-

ylene glycol, pH 4) followed by subsequent reconcentra-

tion of the sample to a final volume of less than 1 mL.

Reduced protein samples were prepared in an inert

atmosphere glove box. An excess of ascorbate ( more than

4.5 equiv; the ascorbate solution was 1.5 mM in deoxy-

genated water) was added to a concentrated sample of

wild-type or mutant TBM, prepared as described above,

that had been deoxygenated by sparging it with water-

saturated argon. Samples were spun down (8,000 rpm,

1 min) to remove any precipitate.

All enzyme samples for XAS contained approximately

20% ethylene glycol. Eighty microliters of each prepared

solution was added to an XAS sample holder via a syringe,

and frozen in liquid nitrogen. The final TBM concentration

of all XAS samples ranged from 150 to 450 lM. Protein

concentrations were determined both by Bradford assays

and by UV–vis (A280) spectroscopy. Molecular weights and

extinctions coefficients (A280) for wild-type and mutant

TBM lacking the His-tag were determined using ExPASy,

on the assumption that all Cys are half-Cys and neglecting

any posttranslational glycosylation (http://www.expasy.org).

The concentrations determined by Bradford assays were

within 5% of the values derived from the absorbance at

280 nm.

Collection and analysis of XAS data

Copper K-edge (8.9 keV) EXAFS and X-ray absorption

near edge structure (XANES) data were collected at the

Stanford Synchrotron Radiation Lightsource operating at

3 GeV with currents between 100 and 80 mA. Cu(I)-con-

taining samples were measured on beam line 9-3 using a

Si[220] monochromator and a rhodium-coated mirror

upstream of the monochromator with a 13 keV energy

cutoff to reject harmonics. A second rhodium mirror

downstream of the monochromator was used to focus the

beam. Data were collected in fluorescence mode using a

high-count-rate Canberra 30-element germanium array

detector with maximum count rates below 120 kHz. A Z-1

nickel oxide filter and a Soller slit assembly were placed in

front of the detector to reduce the elastic scatter peak.

Cu(II) samples were measured on beam line 7-3 using

lower fluxes and unfocused optics, but with a similar beam

line configuration of monochromator and harmonic-rejec-

tion mirror. Energy calibration was achieved by reference

to the first inflection point of a copper foil (8,980.3 eV)

placed between the second and third ionization chambers.

The samples (80 lL) were measured as aqueous glasses

(more than 20% ethylene glycol) at 10–15 K. Six scans of a

sample containing only sample buffer were collected,

averaged, and subtracted from the averaged data for the

protein samples to remove Z-1 Kb fluorescence and pro-

duce a flat pre-edge baseline. Data reduction and back-

ground subtraction were performed using the program

modules of EXAFSPAK [30]. Data from each detector

channel were inspected for glitches, dropouts, or other

nonlinear behavior before inclusion in the final average.

Spectral simulation was carried out using the program

EXCURVE 9.2 [31–34] as previously described [23].

EXAFS data were simulated using a mixed-shell model

consisting of imidazole and S(Met) coordination. First-

shell distances (R) and Debye–Waller factors (2r2) for the

Cu–N(imidazole) and the Cu–S(Met) shell, and the

threshold energy E0 were initially refined. In these pre-

liminary refinements, the imidazole ring outer-shell carbon

and nitrogen atoms were constrained to move relative to

the first-shell Cu–N distance so as to maintain the idealized

ring geometry, and all single- and multiple-scattering

pathways were included in the calculations as previously

described [23]. Later in the refinement, this constraint was

1198 J Biol Inorg Chem (2010) 15:1195–1207

123

lifted, and the outer shells of the imidazole rings were

allowed to float within 10% of their original idealized

positions. In practice, the final outer-shell coordinates for

acceptable fits deviated by less than the permitted amount

from the idealized position. The parameters refined in the

fit included shell occupancy N, copper-scatterer distance R,

and Debye–Waller factor (2r2) for each shell, and the

threshold energy (E0) for photoelectron ionization, which

was constrained to be the same for all shells of scatterers.

Results

XAS of the wild-type enzyme

Previous studies of copper loading to TBM showed that the

enzyme bound 1.9 coppers per protein [9] and sequence

homology between TBM and PHM with respect to copper-

binding residues suggests these two copper centers are

chemically distinct. Since X-ray absorption detects only the

average ligand environment, chemical modeling is required

to gain information specific to each site. In a previous

EXAFS study of PHM, we used the information from the

crystal structure to generate a model which refined the two

copper centers independently [23], the accuracy of which

was later confirmed by Chen et al. [13] using geometry

optimization of the crystal coordinates by density func-

tional theory calculations. In the present case, we applied

similar methods. The data were fit initially by an average

coordination over both copper centers, and were subse-

quently tested for improvements in goodness of fit (F)

when the copper centers were modeled separately. Figure 2

compares the Fourier transforms (FT) and EXAFS spectra

for the oxidized and reduced forms of TBM. The oxidized

form shows a spectrum typical of Cu(II)–imidazole coor-

dination as expected from His ligation. The data fit well to

2.5 His residues and 1.5 O/N ligands, which represent the

average coordination at each copper. No improvement in

the value of F was obtained by modeling the coppers

individually. The EXAFS spectrum of reduced TBM dif-

fers significantly from that of the oxidized protein, showing

a large decrease in amplitude of the first shell in the FT. A

single shell of imidazole groups was generally sufficient to

account for the EXAFS and transform features. Figure 2

also shows the fit of reduced TBM at pH 7.0, corresponding

to two His residues with Cu–N distances of 1.94 A and 0.5

sulfur at 2.24 A (Table 1, reduced protein fit A). The

Debye–Waller term for the Cu–N(His) shell is unusually

large and suggests a large spread in the individual

Cu–N(His) distances. This suggested the presence of two

coordinatively distinct copper centers, and in this case

warranted further exploration of chemical inequivalence

between the copper centers. This was achieved by

assuming inequivalence of the His coordination at each

copper, and simulating the data with two independent

shells of imidazole ligands. Splitting the imidazole scat-

tering into two separate contributions produced no

improvement in the value of the goodness-of-fit parameter

F, but resulted in more reasonable values for the Debye–

Waller factors of each shell (Table 1, reduced protein fit B,

Fig. S1). Notably, one shell of His residues appears to be at

a short distance of 1.88 A, a distance typical of two-

coordinate Cu(I)–imidazole complexes [35–37], whereas

the other shell refines to R = 1.98 A, a distance more

R (Å)

k (Å -1 )-5

0

5

Fo

uri

er T

ran

sfo

rm

0

10

20

30

KeV0.0

0.5

1.0

R (Å)

k (Å -1 )-5

0

5

1 2 3 4 5 6

1 2 3 4 5 6

Fo

uri

er T

ran

sfo

rm

0

10

20

KeV

2 4 6 8 10 12

8.98 9.00

2 4 6 8 10 12

8.98 9.000.0

0.5

1.0

Fig. 2 Phase-corrected Fourier transforms and extended X-ray

absorption fine structure (EXAFS) (upper insets) for oxidized (top)

and ascorbate-reduced (bottom) copper centers in tyramine b-

monooxygenase. Solid black lines are experimental data and dashedred lines are simulations using parameters listed in Table 1. The

lower insets show the X-ray absorption near edge structure (XANES)

region of the spectrum

J Biol Inorg Chem (2010) 15:1195–1207 1199

123

characteristic of three-coordinate Cu(I). We assign the

two- and three-coordinate sites to the H-site and M-site,

respectively.

We also explored fits involving higher imidazole coor-

dination numbers, since homology to PHM predicts that all

five His residues could remain coordinated to Cu(I) in the

reduced form, leading to the predicted average imidazole

per copper of 2.5 [14, 15, 23]. Fits using a single shell of

2.5 His per copper gave acceptable F values but with even

larger Cu–N Debye–Waller factors (2r2 = 0.022 A2,

Table 1, reduced protein fit C). Using the above reasoning,

this implies more heterogeneity between the two Cu(I)

centers, but unexpectedly, two-shell fits with increased

coordination number resulted in extremely poor fits to the

data, with an unacceptably larger Debye–Waller factor at

one copper center. The conclusion that we draw from this

analysis is that the Cu(I) centers are most likely each

coordinated by only two EXAFS-detectable His ligands.

However, given the difficulty in determining EXAFS-

derived coordination numbers to high precision, the

presence of a third His residue at the CuH center cannot be

discounted.

The identity of the remaining ligands at the M-site is

less clear from the EXAFS data. Inclusion of a weak-

interaction 0.5 sulfur donor at 2.24 A improves the fit by

broadening the first shell in the FT sufficiently to fit the

experimental data. However, the Debye–Waller factor is

unusually high for an isolated absorber–scatterer interac-

tion, and suggests there may be two or more conformations

for the Met ligand. Metal-S(Met) coordination often exists

as multiple conformers with the terminal methyl group of

one rotated relative to the other [38]. The best fit to the data

is listed in Table 1, fit A. Figure S1 compares fits using one

versus two groups of imidazoles, while Figure S2 shows

the effect of ommitting the sulfur ligation.

pH dependence of the reduced wild-type protein

PHM shows a pH-dependent structural transition in which

the intensity of the Cu–S interaction increases with

Table 1 Fits obtained to the extended X-ray absorption fine structure of reduced tyramine b-monooxygenase and its Met to His, Asp, and Cys

variants

Cu–N(His)a Cu–O/N Cu–S/Cl -E0

Fb No.c R (A)d DW (A2) No.c R (A)d DW (A2) No.c R (A)d DW (A2)

Oxidized proteins

WT pH 7 0.520 2.5 1.97e 0.012 1.5 1.97e 0.012 4.35

M471H 0.470 2.5 1.97e 0.012 1.5 1.97e 0.012 4.98

M471D 0.513 2.5 1.96e 0.012 1.5 1.96e 0.012 4.35

M471C 0.410 2.5 1.97e 0.011 1.5 1.97e 0.011 4.98

Cu–N(His1)a Cu–N(His2)a Cu–S/Cl

Reduced proteins

WT pH 7 (fit A) 0.328 2 1.94 0.016 0.5 2.24 0.015 2.28

WT pH 7 (fit B) 0.324 1 1.88 0.012 1 1.98 0.007 0.5 2.25 0.014 2.10

WT pH7 (fit C) 0.342 2.5 1.95 0.022 0.5 2.24 0.014 2.28

WT pH 4 0.857f 2 1.94 0.017 1 2.28 0.012 1.88

M471H 0.358 2 1.91 0.011 2.1

M471D 0.359 2 1.91 0.009 1.94

M471C 0.681 2 1.92 0.012 1.93

DW Debye–Waller factor, WT wild typea Fits modeled His coordination by an imidazole ring, which included single- and multiple-scattering contributions from the second-shell (C2/

C5) and third-shell (C3/N4) atoms, respectively. The Cu–N–Cx angles were as follows: Cu–N–C2 126�, Cu–N–C3 -126�, Cu–N–N4 163�, Cu–

N–C5 -163�b F is a least-squares fitting parameter defined as F2 ¼ 1

N

PNi¼1 k6ðData�ModelÞ2

c Coordination numbers are generally considered accurate to ±25%d In any one fit, the statistical error in bond lengths is ±0.005 A. However, when errors due to imperfect background subtraction, phase-shift

calculations, and noise in the data are compounded, the actual error is probably closer to ±0.02 Ae The distances of the Cu–N(His) and Cu–N/O (non-His) shells were constrained to be equalf The larger value of the goodness-of-fit parameter results from data of lower signal to noise

1200 J Biol Inorg Chem (2010) 15:1195–1207

123

decreasing pH with a pK of approximately 4.6 in 2-mor-

pholinoethanesulfonic acid buffer [24]. Given the homol-

ogy between PHM and TBM, it was of interest to

determine whether a similar structural transition was

present in TBM. Figure 3 compares the FTs of reduced

TBM at pH 4 and pH 7, whence it can be seen that the

intensity of the Cu–S peak at R * 2.3 increases dramati-

cally at pH 4. Figure 3 also shows a simulation of the low-

pH data with two Cu–N(His) interactions at 1.94 A and one

Cu–S/Cl interaction at 2.28 A. The increase in shell

occupancy from 0.5 to one Cu–S/Cl suggests that an

additional Cu–S/Cl interaction is observed at low pH but

the correlation between shell occupancy and the Debye–

Waller factor implies that the alternative of a more rigid

(lower Debye–Waller factor) Cu–S interaction should also

be considered. In the PHM system the increase in Cu–S

intensity at low pH could be simulated either by an addi-

tional Cu–S interaction or by allowing the Debye–Waller

factor to decrease at low pH, and it was argued that the

latter would imply a transition from a flexible to a rigid

conformation [24]. The pK for the transition correlated

with the decrease in activity at low pH, and it was sug-

gested that the active form of the enzyme therefore

required a flexible conformation with a weak or fluxional

Cu–S(Met) interaction at CuM. In contrast, the TBM low-

pH data cannot be adequately simulated with 0.5 Cu–S and

a low Debye–Waller factor. Furthermore, the bond length

for the Cu–S/Cl interaction increases from 2.24 A at pH 7

to 2.28 A at pH 4. This increase in bond length is incon-

sistent with a strengthening of the Cu–S bond which is

implied by a lower Debye–Waller factor, and suggests that

the increased intensity more likely arises from an addi-

tional Cu–S/Cl ligation at one of the copper centers at low

pH.

Met to His, Cys, and Asp variants

To probe the role of the essential Met residue at position

471 in TBM, variants with His, Cys, or Asp replacing the

Met were created. Figure 4 compares the spectra of all

three mutants with those of the wild-type protein for the

oxidized proteins. Remarkably, the agreement among all

four experimental spectra is closer than that between the

spectrum of the wild type and its best simulation. There-

fore, we conclude that there is no difference in coordina-

tion between the wild type and the M471X variants,

proving conclusively that residue 471 does not coordinate

the CuM(II) center of the oxidized protein in a manner

detectable by EXAFS spectroscopy. The spectra of the

reduced variants differ significantly from the spectrum of

the wild-type reduced protein (Fig. 5, top), but are all

similar to each other (Fig. 5, bottom), suggesting that

replacement of the M-site residue by His, Cys, or Asp

produces a species with similar coordinate structure. All

three reduced variants can be well fit by an average of two

His residues at each copper with Cu–N(His) distances of

1.91 ± 0.01 A (Fig. 6, Table 1). As expected, the His and

Asp variants show no sign of Cu–S interactions. However,

since the Cys variant exhibits small differences from His

and Asp in the k = 7–9 A-1 energy regime, we tested

whether these arose from a Cu–S contribution of low shell

occupancy, perhaps arising from a small population of

enzyme molecules in a ‘‘Cys-on’’ conformation. When the

Cu–S Debye–Waller factor was fixed at 0.005 A´ 2, a value

typical of Cu(I)–thiolate bonds, the simulations could tol-

erate 14% of a Cu–S conformation, with RCu–S = 2.28 A´

.

However, the goodness of fit was not improved in these

simulations, and the inclusion of the Cu–S wave did not

account for the small differences in EXAFS between

R (Å)0 1 2 3 4 5 6

Fo

uri

er T

ran

sfo

rm

0

10

20

High pHLow pH

R (Å)

k (Å - 1 )-5

0

5

1 2 3 4 5 6

Fou

rier

Tra

nsfo

rm

0

10

20

KeV

2 4 6 8 10 12

8.98 9.000.0

0.5

1.0

Fig. 3 pH dependence of reduced wild-type (WT) tyramine b-

monooxygenase. Top: Comparison of the phase-corrected Fourier

transforms at pH 7 (black) and pH 4 (red). Bottom: Phase-corrected

Fourier transforms and EXAFS (upper inset) for the reduced wild-

type protein at pH 4. Solid black lines are experimental data and

dashed red lines are simulations using parameters listed in Table 1.

The lower inset shows the XANES region of the spectrum

J Biol Inorg Chem (2010) 15:1195–1207 1201

123

k = 7 A-1 and k = 9 A-1. Therefore, the major species

present in all three variants contains only Cu–His coordi-

nation, but we cannot exclude the possibility of a minor

component in M471C having coordinated Cys, which

might possibly account for the observed activity of this

variant. Figure 6 compares experimental and simulated

EXAFS spectra for the His and Cys variants. Simulation

using a two-shell imidazole model resulted in one group of

Cu–His interactions at a short distance of approximately

1.84 A and a second group at a longer distance of

approximately 1.95 A. The separation of 0.11 A is below

the limit of resolution of the experiment given by DR =

p/2Dk equal to 0.15 A. These distances suggest one two-

coordinate and one three-coordinate site, respectively.

Absorption edges

XANES spectra for wild-type TBM and the M471 variants

are shown in Fig. 7 (top). Wild type oxidized and reduced

spectra are compared in Fig. 7 (top). These are typical for

this class of copper monooxygenases, with the oxidized

absorption edge shifted about 7 eV relative to the reduced

protein with a low-intensity 1s ? 3d transition at

8,979 eV, and weak unresolved features on the rising edge

at 8,983.5 and 8,988 eV. The reduced protein exhibits a

resolved feature at 8,983.4 which can be assigned as the

1s ? 4p transition of Cu(I) in a three-coordinate envi-

ronment [37, 39, 40]. All of these XANES features

resemble those of PHM [23] and DBM [10], and confirm

the similarity in coordinate structure of the copper centers

of this class of copper monooxygenases.

Figure 7 (bottom) compares XANES spectra for the

reduced wild-type protein and the three M471 mutant

proteins. The M471 variants also show the 1s ? 4p tran-

sition but it is more intense and shifted slightly to higher

energy (8,984.0 eV). Of particular interest is the almost

exact coincidence of the XANES spectra for all three M471

variants, reinforcing the conclusion from EXAFS analysis

R (Å)

0 1 2 3 4 5 6

Fo

uri

er T

ran

sfo

rm

0

10

20

30

40

WTHisCysAsp

k (Å-1)2 4 6 8 10 12

k3 x E

XA

FS

-10

-5

0

5

10WTHis CysAsp

Fig. 4 Comparison of the experimental EXAFS (top) and phase-

corrected Fourier transform data (bottom) for oxidized WT tyramine

b-monooxygenase and its M-site Met to His, Asp, and Cys variants.

WT black, M471H blue, M471C red, M471D green

k (Å-1)

2 4 6 8 10 12 14

k3 x E

XA

FS

-6

-4

-2

0

2

4

His Cys Asp

R(Å)

0 1 2 3 4 5 6

Fo

uri

er T

ran

sfo

rm

0

5

10

15

20

WTHis

k (Å-1)

4 6 8 10 12

k3 x

EX

AF

S

-4

-2

0

2

4

Fig. 5 Top: Comparison of experimental phase-corrected Fourier

transforms and EXAFS (inset) of ascorbate-reduced WT (black trace)

and the M471 to His variant (red trace) of tyramine b-monooxygen-

ase. Bottom: Comparison of the experimental EXAFS data of the Met

to His (blue), Met to Cys (red), and Met to Asp (green) variants

1202 J Biol Inorg Chem (2010) 15:1195–1207

123

that they all have very similar coordination at both copper

centers. The increased intensity indicates that the average

coordination number has decreased, and is consistent with

the observed decrease in the Cu–N(His) distance, which

drops from 1.94 A in the reduced wild type to 1.91 A in the

mutant proteins. In general, Cu(I)–N(His) distances corre-

late with the coordination number, with the distance

decreasing as the coordination number drops. The decrease

in coordination can be explained by assuming that residue

471 no longer coordinates at the M-site, and that the latter

is coordinated by just two His residues. Alternatively, the

loss of M471 may also induce changes at the H-site as

recently observed in the crystal structure of the M314I

mutant of PHM [25].

Discussion

TBM shares structural features with PHM and DHM.

Sequence alignment (Fig. 8) shows that all the residues

known to be involved in copper binding in PHM are con-

served in TBM, and suggests that TBM likewise contains a

pair of mononuclear copper centers bound by H247, H248,

and H317 (H-site) and H396, H398, and M471 (M-site).

However, kinetic studies with TBM have revealed several

important differences between the mechanism of the insect

enzyme and the established mechanism of the mammalian

homologues, with respect to the interaction of the enzymes

with substrate and ascorbate [29]. We used XAS to

investigate the coordination environment of the copper

centers of TBM with the purpose of understanding the

structural origins of these mechanistic differences. Data on

the oxidized and reduced forms of the wild-type protein

bear a close resemblance to those from PHM establishing a

homology with respect to copper binding. Thus, we con-

clude that in oxidized TBM Cu(II) is bound by three His

residues and a water molecule at the H-site, and by two His

R (Å)

-1 )

2 4 6 8 10 12-5

0

5

1 2 3 4 5 6

Fou

rier

Tra

nsfo

rm

0

10

20

KeV

8.98 9.000.0

0.5

1.0

Met to His

R (Å)

k (Å

k (Å -1 )

2 4 6 8 10 12-5

0

5

1 2 3 4 5 6

Fou

rier

Tra

nsfo

rm

0

10

20

KeV

8.98 9.000.0

0.5

1.0

Met to Cys

Fig. 6 Phase-corrected Fourier transforms and EXAFS (upper insets)

for reduced M471 to His (top) and reduced M471 to Cys (bottom)

variants of tyramine b-monooxygenase. Solid black lines are exper-

imental data and dashed red lines are simulations using parameters

listed in Table 1. The lower insets show the XANES region of the

spectrum

keV8970 8980 8990 9000

F/I 0

0.0

0.5

1.0

WT His Cys Asp

keV

8970 8980 8990 9000

F/I 0

0.0

0.5

1.0OxidizedReduced

Fig. 7 Comparison of XANES of the copper centers of tyramine

b-monooxygenase. Top: Oxidized versus reduced wild-type proteins.

Bottom: Reduced proteins: WT black, Met to His blue, Met to Cys

red, Met to Asp green

J Biol Inorg Chem (2010) 15:1195–1207 1203

123

residues and two solvent molecules at the M-site. On

reduction, the solvent coordination is lost, and M471

moves closer to CuM to form a three-coordinate center

comprising H396, H398, and M471. Although the EXAFS

data are best fit to two His residues at each copper site, the

similarity of the spectra to previous data on reduced wild-

type PHM and DBM, and the additional crystallographic

information available for PHM, supports a three-coordinate

CuH site, with one weakly bound His residue undetectable

by EXAFS. Thus, loss of solvent at the H-site appears to

result in a three-coordinate Cu(I) center in the reduced

form of wild-type TBM, bound by H247, H248, and H317.

Thus, mechanistic differences among the three enzymes do

not appear to result from an altered copper coordination

environment in TBM.

The pH dependence of the EXAFS spectrum of reduced

wild-type TBM shows an increase in intensity of the Cu–S

interaction similar to that reported previously for PHM but

with important differences [24]. Although the pKa for the

pH transition was not determined, it is likely that a similar

Met-on to Met-off transition controls the catalytic activity

of TBM at low pH. In PHM, the Met-off form was shown

to be catalytically active, whereas the Met-on form was

inactive, and it was proposed that Met-off represented the

weakly bound, fluxional CuM–S(Met) form fully described

by EXAFS and crystallography. It was proposed [24] that

Met fluxionality provided the necessary protein dynamics

to bring the enzyme into the transient configuration critical

for hydrogen tunneling. The identity of the Met-on form is

less clear, but two possibilities exist: (1) a rigid, strongly

bound CuM–S(Met) lacking in some essential fluxional

component necessary for catalysis, or (2) a Cu–S interac-

tion formed between either a copper center and a different

sulfur-containing residue or, alternatively, reaction of one

of the copper centers with an exogenous chloride ion. The

fact that TBM low-pH data fit better to one Cu–S/Cl per

TBM with a longer bond length yet smaller Debye–Waller

factor is suggestive of the latter conclusion. All of the ten

Cys residues in PHM are present as disulfides and it is

therefore unlikely that additional Cu–S coordination

involves a Cu(I)–Cys(thiolate) interaction. However, the

strong preference of Cu(I) centers for Met coordination, for

example in the periplasmic copper binding proteins CusF

[38], CusB [41], pcoC [42], and CopK [43], indicates that a

Met residue is a likely candidate. PHM and TBM share

only two conserved Met residues, the catalytic-site M314/

M471 residue and M109/M249, which is contiguous to the

H-site copper-binding His residues, but is rotated away

from the copper on the opposite side of the b-sheet. If the

Met-on form represents copper coordination by a different

Met residue, then CuH coordination to M109/M249 seems

a likely option. This would require a significant confor-

mational change induced by protonation of an as yet

unassigned residue. The resultant loss of activity could be

the result of a change in reduction potential of the H-site

expected if a Met residue replaces a His residue. Alterna-

tively, the reorientation of b-structure required to bring

M109/M249 into coordinating distance of the CuH center

would most likely disrupt the hydrogen bond between

H108 and E170 (numbering refers to PHM) which has been

proposed as part of the putative substrate-mediated electron

transfer pathway between the two copper centers [15].

Fig. 8 Sequence alignment

using Clustal W 2.0.10 of

members of the family of

mononuclear monooxygenases.

Sequences for peptidylglycine

monooxygenase (PHM) and

dopamine b-monooxygenase

(DBM) are from rat, for

tyramine b-monooxygenase

(TBM) are from Drosophilamelanogaster, and for

monooxygenase X (MOX) are

from human. Metal-binding

residues and conserved Met

residues are shown in red

1204 J Biol Inorg Chem (2010) 15:1195–1207

123

The possibility that the additional interaction is due to

binding of exogenous chloride at low pH cannot be

excluded on the basis of EXAFS analysis, but seems less

likely from a chemical perspective. There is no reason why

chloride binding should be pH-dependent as HCl is a

strong acid. Additionally, similar chemistry has recently

been observed with PHM in samples which do not contain

chloride ion [24]. Therefore, although we cannot exclude

chloride, coordination by M249 seems the most likely

option.

The unique coordination of the copper centers has been

shown in PHM and DBM to induce novel monooxygenase

chemistry. In these sister proteins, catalysis begins by

coordination of an oxygen molecule at the reduced

mononuclear CuM center as observed crystallographically

in a precatalytic complex with a slow substrate [16]. The

O–O bond length in this complex is consistent with a

Cu(II)–superoxo species, and dovetails with other bio-

chemical [17, 19] and theoretical [3, 20] studies that

implicate Cu(II)–O2�- as the reactive oxygen intermediate.

The electrophilic nature of such an intermediate is expected

to lead to efficient hydrogen-atom abstraction from an a-

carbon of peptide substrate [44] or the benzylic position of

phenethylamines, and calculations suggest a preference for

side-on superoxide coordination [3, 20]. However, among

the growing number of mononuclear Cu(II)–superoxo

model complexes [26], only two are reactive toward

hydrocarbon substrates [45, 46], and these exhibit end-on

superoxo coordination. A more common reactivity of

mononuclear Cu(II)–superoxo species is dimerization with

another molecule of the Cu(I) parent complex to generate

the dinuclear Cu(II)–peroxo complex. The large spatial

separation of the copper centers in PHM and by analogy

TBM prevents this chemistry, and thus allows the potent

electrophilic reactivity of the mononuclear Cu(II)–superoxo

species to be fully expressed in the form of hydrogen-atom

abstraction from the substrate to form a [CuM(II)-OOH]

(peroxo) intermediate. Subsequent steps in the reaction

pathway are less clear and alternative mechanisms have

been proposed that involve long-range electron transfer

from CuH either before [17] or after [20] the transfer of an

OH group to the substrate radical.

An unresolved issue relating to this mechanism is the

role of the M-site Met residue in stimulating catalysis. As

stated earlier, the flexibility of the Met ligand has been

invoked as a necessary element for catalysis. In addition,

the Met ligand is believed to influence the Cu(II)/(I) redox

equilibrium at the CuM site [17] and thus affect the extent

of dioxygen activation. A preference for formation of an

initial Cu(I)–O2 versus a Cu(II)–superoxide species is

thought to stem from the effect of sulfur ligation. This

equilibrium shift in favor of an unactivated bound dioxy-

gen species would greatly reduce uncoupling and the

formation of harmful reactive oxygen species, prior to C–H

activation. The alternative residues Asp, His, and Cys, in

place of the Met ligand at CuM, were thus chosen to further

examine the significance of this residue.

Mutation of M471 in TBM to His, Asp, or Cys resulted

in a large decrease in activity, with only the M471C variant

displaying measurable activity, albeit coincident with

enzyme inactivation [9]. In the case of PHM, the M314X

variants had no detectable activity in assays of spent media

from Chinese hamster ovary cell overexpression [21, 22].

One of the goals of the present work was to provide a

structural basis for the reduction/loss in catalytic activity

by examining the EXAFS-derived structures of the M471X

(X is His, Cys, Asp) variants. Results on the oxidized

protein establish that the Met ligand does not bind strongly

to the Cu(II)M center, as no Cu–S interaction is observed.

This result is similar to those for PHM and DBM, where no

Cu–S EXAFS interaction is seen in the Cu(II) form [10,

23], owing to a long axial Cu–S(Met) interaction [13], and

leads to the prediction that His and Asp would likewise

bind axially and be undetectable by EXAFS, as is observed

experimentally. The Cys variant might have been expected

to show some properties of type 1 copper centers due to the

N2S coordination, but it too appears to produce no per-

turbation on the wild-type spectrum with no observable

thiolate-to-Cu(II) charge transfer. The EXAFS results

together with earlier EPR data [9] therefore suggest that the

oxidized CuM site accommodates all four side chains

without significant structural rearrangement.

In contrast, the reduced forms of wild-type and variant

TBM show significant differences. At pHs at or above the

activity maximum (pH 5.5–6), reduced wild-type TBM

exhibits a Cu–S interaction equivalent to 0.5 Cu–S at

2.25 A with a Debye–Waller factor (2r2 = 0.015 A2)

which is high for a strongly coordinated ligand, and may

suggest some fluxional behavior involving either multiple

Met conformations (as is seen, for example, in the peri-

plasmic chaperone CusF [38]), or alternatively rapid

coordination and dissociation of the thioether moiety.

Such behavior has been observed in Cu(I)NXS(thioether)

models of the CuM site including pendant thioether con-

taining b-diketiminate Cu(I) complexes which show

temperature-dependent changes in the thioether-bonded

methylene proton NMR resonances [28], and in TMPA-

derived (TMPA = tris(2-pyridylmethyl)amine) N3S(thio-

ether) models where the thioether arm is easily displaced

by CO [47]. Cu–S(Met) fluxionality may be an important

feature of the M active site, either to provide facile

conversion to axially coordinated Cu(II)–S(Met) upon

oxygen binding and formation of the Cu(II)–superoxo

intermediate [13], or to provide kinematic coupling to a

normal mode critical to hydrogen tunneling during the

hydrogen-atom abstraction step [1, 24, 48].

J Biol Inorg Chem (2010) 15:1195–1207 1205

123

Comparison of the EXAFS data for the His, Asp, and

Cys variants with EXAFS data for reduced wild-type

TBM shows differences, but analysis leads to the con-

clusion that none of these substitutions lead to detectable

coordination to copper. The presence of both CuH and

CuM makes it difficult to isolate the structural changes at

the M-site, and the data give only the average change at

both coppers. However, since the H-site locus is

unchanged in these variants, it is reasonable to assume

that the observed structural changes derive mainly from

the M-site substitutions. Analysis of the EXAFS data

gives a similar structure for all three variants, with loss of

the Cu–S component, and decrease in the residual two

Cu–N(His) bond lengths. In parallel, the XANES data

shows an increase in intensity of the 8,983-eV edge fea-

ture. These changes are consistent with a decrease in

coordination number and suggest a simple interpretation

that the M-site becomes two-coordinate. Although coor-

dination of oxygen-donor groups, such as Asp, is less

common among copper proteins, it is surprising that

neither His nor Cys seems capable of significant binding

to CuM. The fluxional Cu(I)–Met interaction, and its

movement by some 0.5 A on copper oxidation indicates

the absence of steric restrictions in the site, and modeling

confirms this conclusion (Fig. 1c). In other systems, Met

to His substitution appears quite facile, for example, the

M121H variant of Alcaligenes denitrificans azurin, where

despite the ‘‘rack-induced’’ rigidity of the cupredoxin

fold, H121 moves closer to the copper and becomes a

fourth strongly bound ligand [49]. Likewise, recent work

from one of the authors’ laboratories (N.J.B.) has dem-

onstrated Met coordination in the H135 to Met substitu-

tions of Bacillus subtilis Sco (unpublished).

The crystal structure of the PHM M314I variant sug-

gests one possible reason for the failure of the alternative

His or Cys residues to coordinate [25]. Here the CuM site in

oxidized M314I PHM accommodates the mutation by

replacing M314Sc with a water molecule, and by shifting

the positions of the other coordinating residues (H242,

H244, and a water molecule), to form a distorted tetrahe-

dron. I314 is rotated away from the copper in a different

conformation. Interestingly the PHM M314I structure also

shows significant perturbation at the H-site. H107 and

H172 are now linearly coordinated with H108 dissociated.

The M471X substitutions in TBM may favor this altered

conformation. Therefore, one hypothesis for the essential

role of Met at the M-site is that it uniquely stabilizes the

protein in its catalytic conformation in addition to, or

instead of, electronic tuning of the reactivity of the Cu(II)–

superoxo [17] and/or the Cu(II)–oxyl intermediates [20] as

previously proposed.

This does not, however, explain the singular activity of

the M471C TBM mutant. The lack of sulfur-atom

coordination by the Cys residue to CuM(I) does not pre-

clude substrate turnover. The observed activity of the

M471C mutant may arise from a small population of

enzyme in a Cys-bound conformation, which is not

inconsistent with the EXAFS analysis. Low occupancy of

the Cys-bound form could also provide an explanation for

the inactivation of M471C TBM during the reaction with

substrate. According to the current hypothesis, the fluxional

dynamics of the Cu–Met bond prevent uncoupling of

dioxygen and C–H activation during the catalytic cycle.

Since a fully liganded Cys may be expected to modulate

the chemistry at the CuM site as efficiently as the Met

ligand, the inactivation of M471C via oxidative damage

(R.L. Osborne and J.P. Klinman, unpublished data) may

be a direct consequence of its poor coordination to the

reduced metal. Further work is needed to evaluate these

aspects.

Acknowledgments We thank Andrew Bauman for help with col-

lection of XAS data. We gratefully acknowledge the use of facilities

at the Stanford Synchrotron Radiation Lightsource, which is sup-

ported by the National Institutes of Health Biomedical Research and

Technology Program Division of Research Resources, and by the US

Department of Energy Office of Biological and Environmental

Research. The work was supported by grants from the National

Institutes of Health (R01 NS027583 to N.J.B, R01 GM0257651 to

J.P.K., and GM067351 to C.H.).

Open Access This article is distributed under the terms of the

Creative Commons Attribution Noncommercial License which per-

mits any noncommercial use, distribution, and reproduction in any

medium, provided the original author(s) and source are credited.

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